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1
PEROXISOMAL PLANT 3-KETOACYL-COA THIOLASES’ STRUCTURE AND ACTIVITY
ARE REGULATED BY A SENSITIVE REDOX SWITCH*
Valerie E. Pye1,3,4
, Caspar E. Christensen1, James H. Dyer
2, Susan Arent
1,5, Anette Henriksen
1,3
From the Protein Chemistry Group, Carlsberg Laboratory, Gamle Carlsberg Vej 10, DK-2500 Valby,
Denmark1 and the Department of Chemistry and Biochemistry, Montclair State University, Upper
Montclair, NJ 07043 USA2.
Running title: Redox control of peroxisomal thiolase. 3Address correspondence to: Anette Henriksen, Protein Chemistry Group, Carlsberg Laboratory, Gamle
Carlsberg Vej 10, DK-2500 Valby, Denmark. Fax: +45 33274708 E-mail: [email protected], and Valerie
Pye, School of Biochemistry and Immunology, Trinity College Dublin, Dublin 2, Ireland. Fax:
+353(0)18964257 E-mail: [email protected].
The breakdown of fatty acids, performed by
the -oxidation cycle, is crucial for plant
germination and sustainability. -oxidation
involves four enzymatic reactions; the final
step, where a two carbon unit is cleaved from
the fatty acid, is performed by a 3-ketoacyl-
CoA thiolase (KAT). The shortened fatty acid
may then pass through the cycle again (until
reaching acetoacetyl-CoA) or be directed to a
different cellular function. Crystal structures of
KAT from Arabidopsis thaliana and Helianthus
annuus have been solved to 1.5 Å and 1.8 Å
resolution respectively. The dimeric structures
are very similar and exhibit a typical thiolase
fold; dimer formation and active site
conformation appear in an open, active,
reduced state. Using an interdisciplinary
approach, we confirm the potential of plant
KATs to be regulated by the redox
environment in the peroxisome within a
physiological range. In addition, co-
immunoprecipitation studies suggest an
interaction between KAT and the
multifunctional protein, which is responsible
for the preceding two steps in -oxidation, that
would allow a route for substrate channeling;
we suggest a model for this complex based on
the bacterial system. Fatty acids are fundamental biomolecules
that are abundant in all life forms. With their
enormous variation in chain length and degree of
saturation they are essential for energy storage,
form structural entities in bio-membranes and
serve as signaling molecules. Fatty acids are
broken down in a cyclic manner, by two carbons
at a time, to generate a range of products by the
process known as -oxidation (1). In higher plants
(2) and yeast (3) -oxidation of all forms of fatty
acid occurs in the peroxisomes. In plants, -
oxidation is essential for a plethora of
physiological roles including responses to
senescence and starvation, fatty acid turnover and
regulation of plant lipid composition. Germinating
seeds depend on -oxidation for the mobilization
and release of energy stored in the seed (4).
Arabidopsis thaliana seeds deficient in -
oxidation enzymes are unable to germinate
without an external sugar source; they have large
and unusual peroxisomes and accumulate C16-
C20 fatty acids (5-6). -oxidation is also
responsible for the synthesis of jasmonic acid (JA)
(7) and indole-3-acetic acid (auxin, IAA), via
conversion from indole-3-butyric acid (IBA) (8),
which serve as crucial plant hormones regulating
plant development and responses to biotic and
abiotic stress. Hydrogen peroxide, produced as a
by-product during -oxidation, is used by catalase
to oxidize different toxins (e.g. alcohols) and plays
an important role in cellular signaling (9).
-oxidation is composed of four reactions:
Firstly, the CoA-activated acyl chain is oxidized to
2-trans-enoyl-CoA by an acyl-CoA oxidase
(ACX) (10) producing H2O2 as a by-product. The
double bond is then reduced by 2-trans-enoyl-
hydratase forming L-3-hydroxyacyl-CoA,
followed by oxidation by NAD+ dependent L-3-
hydroxyacyl-CoA dehydrogenase. Both the
hydratase and dehydrogenase activities are
performed by a multifunctional protein (MFP)
(11-12). 3-ketoacyl-CoA thiolase (KAT) performs
the last step of -oxidation, thiolytically cleaving a
two carbon unit from 3-ketoacyl-CoA (13). The
shortened fatty acyl-CoA can then be subjected to
further rounds of -oxidation or can be directed to
other pathways. In mammals there are two distinct
pathways for -oxidation: very long chain fatty
acids and bile acid intermediates are shortened in
peroxisomes, while medium to long chain mono-
and di-carboxylic fatty acids, methyl-branched
fatty acids and prostaglandins are broken down in
mitochondria (14). In both mitochondrial and
http://www.jbc.org/cgi/doi/10.1074/jbc.M110.106013The latest version is at JBC Papers in Press. Published on May 12, 2010 as Manuscript M110.106013
Copyright 2010 by The American Society for Biochemistry and Molecular Biology, Inc.
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prokaryotic -oxidation systems tri-functional
multi-enzyme complexes (comprising MFP and
KAT activities) exist, allowing for metabolite
channeling (15-18). The crystal structure of the
Pseudomonas fragi tri-functional-enzyme complex
is known and consists of two -subunits (hosting
MFP functions and from now on referred to as
PfMFP) and two -subunits (hosting KAT
functions and from now on referred to as PfKAT)
in a dynamic complex (16). Plants contain a
number of isozymes for each step in -oxidation;
the ACXs are best defined with regards to having
different but overlapping substrate specificities.
Communication between the isoforms via a
protein-protein complex, similar to that found in
Pseudomonas fragi, could direct and control the
flow of metabolites through -oxidation, but to-
date no such complex in plants has been reported.
There are two distinct forms of 3-
ketoacyl-CoA thiolases. Type I is 3-ketoacyl-CoA
thiolase (KAT; EC 2.3.1.16), a catabolic enzyme
performing the reverse Claisen condensation
reaction involved in e.g. the -oxidation cycle.
The type II thiolase is acetoacetyl-CoA thiolase
(ACAT; EC 2.3.1.9) and is involved in the
anabolic mevalonate pathway performing Claisen
condensation. Arabidopsis has been shown to
possess three KAT genes encoding three
peroxisomal KAT isoforms (1, 2 and 5.2) and one
cytosolic, lacking the N-terminal 43 residues, (5.1)
and two ACAT genes encoding four cytosolic
ACAT isoforms (1.1, 1.2, 2.1 and 2.2) and one
peroxisomal (1.3) (13). In Arabidopsis, KAT2 is
the major type I thiolase isoform present during
germination and seems to be important for TAG
synthesis during seed development as well (6).
KAT1 and KAT5 are not abundant in seedlings
and their roles are less well defined, KAT5 is
possibly involved in flavonoid biosynthesis (13).
KAT2 is reported to be active for the full range of
fatty acid lengths, from acetoacetyl-CoA upwards
(6) and thus, in contrast to the first step involving
substrate-specific ACXs, the last step of -
oxidation is seemingly unspecific. The first
structure of KAT was solved from Saccharomyces
cerevisiae (ScKAT) (19-20). This structure has a
typical thiolase-like fold, is a homodimer, and
presumably in an active state. The structure of
KAT2 from Arabidopsis (AtKAT2) was solved
later, and also shows a homodimer with a thiolase-
like fold (21). However, the AtKAT2 structure was
noticeably different to the ScKAT structure as a
disulphide bridge had formed involving the
presumed active site nucleophile, causing the
active site to be inaccessible to substrate and
resulted in a change in the loop structure that caps
the active site and a different dimer arrangement.
A regulation of -oxidation by redox potential was
suggested, however, the authors did not produce
either a structure of AtKAT2 in an active, reduced
state or results suggesting oxidation to take place
within a biologically relevant redox range.
Recently, a human peroxisomal KAT (HsKAT)
structure has been deposited in the PDB database
by SGC (PDB accession 2IIK), this is also in the
active, reduced state.
Peroxisomes provide a challenging
environment for enzyme stability. The redox
environment of the peroxisome was traditionally
considered to be stable and reducing due to the
presence of catalase and its role in the breakdown
of H2O2 derived from photorespiration, oxidative
processes and the decomposition of radical oxygen
species (22-23). However, H2O2 is now recognized
as an important signaling molecule in both plants
and animals (for a review see (9)), and plant
peroxisomes have been shown to be net producers
of H2O2, releasing it to the cytosol ((24-25) and for
a review see (26)). The sub-cellular distribution of
H2O2 is unknown, but the total concentration of
H2O2 in unstressed leaves has been measured to be
as high as 3.6 μmol/g (fresh weight) translating to
an average of 3.6 mM (27-28). The findings that
the diene containing fluorescent probe BODIPY-
PTS1 is oxidized in the peroxisomes of rat
fibroblasts upon incubation with eicosanoic and
phytanic acids suggest that the redox state of this
organelle is unstable despite the redox buffering
effects of the anti-oxidant defence systems present
in peroxisomes (29). Stressing the need for
regulation even further, catalase itself is inhibited
at prolonged exposure to H2O2 generated at 2
nmol/(ml min) particularly in the absence of
NADPH (30-34). Additionally, peroxisomes have
been shown to change their morphology
dramatically in response to H2O2 stresses (35). The
inactivation of catalase by H2O2 would lead to a
breakdown of the peroxisomal antioxidant defense
system giving rise to toxic conditions, oxidative
damage and eventually cell death. Therefore a
stable and balanced intracellular redox
environment is crucial for all cells; a control point
in the -oxidation cycle would aid the control of
the redox conditions in the peroxisome.
Here we present results demonstrating that
the redox state and activity of AtKAT2 are
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affected at biologically relevant conditions, and
crystal structures of two plant thiolases: AtKAT2
and sunflower (Helianthus annuus, HaKAT) in a
reduced, open, active conformation to high
resolution. We discuss the potential redox control
of -oxidation and initiated from our co-
immunoprecipitation pull-down assays we suggest
a possible complex formation with MFP. For
completeness we have included in the
supplemental material a comparison of the KAT
structures presented here with other available
peroxisomal type I KAT structures at a structural
and substrate binding level.
EXPERIMENTAL PROCEDURES
Cloning, expression and protein purification - The
full-length HaKAT2 construct (36) was recloned
to remove the N-terminal signaling sequence and
insert a C-terminal His6-tag. The construct for
HaKAT2 aa17-449 was made using Forward 5'-G
GAATTCCATATGCCCTCTTCTACTTCTTCA-
TCCTTGG-3’ and Reverse 5'-GAATTGCGGCG-
TAGTGATGGTGATGGTGATGTTTAGCTTA-
ATTGCAC-3’ primers. The resulting PCR-
fragment was ligated into a pSTblue-1 vector
using the Perfectly Blunt Ligation kit (Novagen).
The purified plasmid was cut with restriction
enzymes NdeI and NotI (New England Biolabs)
and these sites were used for inserting the PCR
product into the pET24b expression vector
(Novagen), followed by transformation of super
competent XL1-blue cells (Stratagene). Protein
expression was carried out using BL21(DE3) cells.
An expression construct encoding AtKAT2 aa36-
462 with an N-terminal enterokinase excisable
His6-tag was prepared in pET46 Ek/LIC from
clone pda03502 (RIKEN) using LIC technology
(Novagen) and the primers 5’-GACGACGAC-
AAGATGGCTGGGGACAGTGCTGCATAT-3’
and 5’-GAGGAGAAGCCCGGTCAGGCGTCC-
TTGGACAA-3’. Successful cloning was verified
by colony PCR and sequencing, and the construct
was used to transform RosettaGami2(DE3) cells
by electroporation. Both proteins were expressed
and purified in the same manner. 1l 2 x YT media,
supplemented with appropriate antibiotics, was
inoculated with a 1/50 overnight culture and
grown to an OD600 = 0.5 at 37oC. Cells were then
cooled to 25oC before inducing with 1 mM IPTG
and incubated (shaking) at 25oC for 18 h. Cell
pellets were resuspended in a pre-chilled solution
of 50 mM Tris(HCl), 200 mM NaCl pH 7.7 and
lysed with bugbuster (Sigma; 1/10 dilution) and
benzonase nuclease (Novagen; 1/1000 dilution)
for 1 h at room temperature. The lysed cell pellets
were centrifuged at 18000 g for 1 h and the
supernatant applied to a 5 ml Ni2+
affinity column
(Amersham Pharmacia). After washing, the
protein was eluted by an imidazole gradient.
Protein fractions were assessed by SDS-PAGE,
concentrated using a Vivaspin20, 30000 MWCO
filter device (Sartorius) and applied to a Hiload
26/60 Superdex 200 prep grade (Amersham
Pharmacia) size exclusion column pre-equilibrated
and run with 50 mM Tris(HCl), 200 mM NaCl pH
7.7 at 4oC. Protein fractions were again assessed
by SDS-PAGE and then dialysed overnight
against 20 mM Tris(HCl), 150 mM NaCl, 2 mM
EDTA, 5% (v/v) glycerol, 2 mM DTT, pH 9.0.
Thiolase activity assay - The activity of AtKAT2
was determined spectroscopically, by monitoring
the absorbance increase at 232 nm due to the
formation of new acyl-CoA ester bonds (37). All
buffers were deoxygenated by purging in an
anaerobic glovebox for 1 h. The reactions
consisted of 50 µM acetoacetyl-CoA and 100 µM
CoA in 35 mM Tris-HCl, 100 mM NaCl, pH 8.5
and were started by the addition of 5 nM AtKAT2
pre-incubated with 20 mM cysteamine (CSH;
Sigma) or 10 mM cystamine (CSSC; Sigma) at
25oC. The enzyme preparations were diluted 2500
times before sampling resulting in a final
concentration of CSH and CSSC of 8 and 4 μM
respectively. Absorbance change at 232 nm was
recorded in a quartz micro-cuvette with a lambda-
45 UV-vis spectrophotometer equipped with a
peltier thermal control element set to 25oC (Perkin
Elmer Inc., USA). Background reactions
containing the buffer and CSH or CSSC were
recorded and used to normalize the data. The half-
lives were determined by fitting the equation for
exponential decay to the data. Acetoacetyl-CoA
and CoA were prepared immediately before
mixing and concentrations checked by UV-
spectroscopy using 257 = 16.840 per M cm for
CoA, 260 = 16.000 per M cm for acetoacetyl-CoA
with an 260/232 = 1.89 (38).
Determination of reduction potential of AtKAT2 -
Tryptophan fluorescence of 5 M AtKAT2 in 35
mM Tris-HCl, 100 mM NaCl, pH 8.5 was
recorded after 15 min. incubation at 25oC with
varying ratios of CSH to CSSC keeping the total
concentration of sulfur constant. 50 l reactions
were prepared in half-area UV 96-well microplates
(Corning 3679, Corning Incorporated, New York)
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and wavelength scans recorded in a fluorescence
platereader (SpectraMAX EM/Gemini, Molecular
Devices, California) exciting at 285 nm and
reading emission from 325-450 nm with 325 nm
cut-off and photomultiplier sensitivity set to
medium. Control experiments with 4 mM H2O2 or
1 mM DTT were included. The emission at 333
nm was plotted against [CSH]2/[CSSC] and the
midpoint of transition found to be 0.17 M by
fitting a two-state function with variable slopes to
the data (Eq. 1).
TR
cxmEXP
TR
cxmEXPxx
xf)(
1
)()(()(
)(%50
%50
1100 Eq. 1.
n and n represent the y-axis intercept and slope
of the first (n=0) and second (n=1) linear parts of
the curve, R is the gas constant, T is the
temperature and c50% and m represents the
midpoint of transition and its slope, respectively.
The midpoint of transition was determined and
used with the Nernst equation and a standard
electrode potential of CSH of -260 mV (39) to
estimate the midpoint electrode potential of
AtKAT2. It was found to be -372 mV at pH 8.5
and -283 mV at pH 7.0. The electrode potential of
AtKAT2 was assumed to follow the -59 mV per
pH unit correlation previously identified for
thioredoxins (40-41). Far UV-CD spectra of 5 M
AtKAT2 incubated with 10 mM CSH or 5 mM
CSSC were recorded in the same buffer as above
at 25oC in a JASCO J-810 CD-spectrometer with a
1 mm light-path quartz cuvette, at 20 nm/min, 0.1
nm pitch and averaging 6 scans.
Crystallisation - HaKAT at 10 mg/ml in 20 mM
Tris(HCl), 150 mM NaCl, 2 mM EDTA, 5% (v/v)
glycerol, 2 mM DTT, pH 9.0 was mixed in a 1:1
ratio with 20% (w/v) PEG 3350, 0.2 M
Li(CH3COO)2, 0.002% (v/v) ethylene glycol and
crystallized by vapour diffusion (hanging drop)
against a reservoir containing 20% (w/v) PEG
3350, 0.2 M Li(CH3COO)2, 0.01 M taurine.
Crystals were flash frozen in liquid nitrogen using
30% (v/v) ethylene glycol as cryo-protectant.
AtKAT2 at 11.8 mg/ml in 20 mM Tris(HCl), 150
mM NaCl, 2 mM EDTA, 5% (v/v) glycerol, 2 mM
DTT, pH 9.0 was mixed in a 1:1 ratio with 20%
(w/v) PEG 3350, 0.2 M Li(CH3COO)2, 0.002%
(v/v) ethylene glycol, 2 mM C12-CoA and
crystallized by vapour diffusion (hanging drop)
against a reservoir containing 20% (w/v) PEG
3350, 0.2 M Li(CH3COO)2. Crystals were flash
frozen in liquid nitrogen using 30% (v/v) ethylene
glycol as cryo-protectant. In both cases crystals
grew within one week.
X-ray data collection, structure solution and
refinement - X-ray data collection and refinement
details are given in Table 1. 5% of reflections
flagged for Rfree were chosen in resolution shells
using SFTOOL because of assumed non-
crystallographic symmetry (NCS). A preliminary
HaKAT data set was collected in-house to 2.5 Å
resolution (data not shown), the AtKAT2 (21)
monomer was used as a search model for 2
molecules per asymmetric unit using PHASER
(42). One clear solution resulted; this structure
was used for refinement against the higher
resolution HaKAT2, data reported here, (the same
Rfree set was applied and extended). Initial rigid
body refinement was performed, treating each
monomer as a rigid body. Cycles of restrained
refinement including motions for
translation/libration/screw (TLS) (4 segments per
chain, defined using TLS server (43) and selected
on B-factor distribution; resulting in 1 group for
the N-terminal domain, 2 groups for L-domain and
1 group for C-terminal domain) in Refmac (44-
45), the use of TLS did lower the Rfree and Rwork
values. Medium NCS restraints (chain A and B)
were used in initial rounds of refinement. These
were loosened for the rest of the refinement; main
deviations between the molecules occur in the
positioning of the -hairpin in the L-domain.
Water molecule assignment and manual rebuilding
was performed in COOT (46) It was noticed that
the B-factors assigned to the water molecules
during this TLS refinement were largely
unexpected, this was also reflected in the electron
density, with B factors either suspiciously high
with positive density or suspiciously low with
negative density. This is reported as a possible bug
in the current version of Refmac (v 5.5.0035).
Therefore, the final rounds of refinement were
performed in PHENIX (47), using the default
settings, and resulted in more realistic B-factors
and density for the water molecules. The HaKAT
dimer model was used as the search model for the
AtKAT2 data using PHASER and resulted in one
clear solution. Refinement proceeded as per the
HaKAT structure. PDB co-ordinates and structure
factors have been deposited with the PDBe
databank with accession codes 2WUA and 2WU9.
Co-Immunoprecipitation Studies - Enriched
peroxisomal lysate was isolated in the following
manner: Seedlings (Brassica napus cv.) were
grown in the dark for 7 days, the cotyledons were
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then harvested, homogenized in 10 ml 0.25 M
sucrose, 0.1 M Tris(HCl) pH 7.4, 1 mM EDTA, 1
mM MgCl2, 0.1% ethanol, complete EDTA free
protease inhibitor (Roche) and filtered through 2
layers of myra-cloth. Cell debris was removed by
centrifugation, 600 g for 5 mins. Peroxisomes
were sedimented by centrifugation (15,000 g, 20
mins) and resuspended in 1 ml 0.1 M Tris(HCl)
pH 7.4, 1 mM EDTA, 1 mM MgCl2, 150 mM
NaCl, complete EDTA free protease inhibitor
(Roche) and incubated on ice for 20 mins. to lyse
the peroxisomal membranes. The lysate was
cleared twice by centrifugation at 15,000 g for 20
mins. The cleared lysate was then used for co-
immunoprecipitation studies using the Seize X
protein A kit (Pierce), following manufactures
guidelines, immobilizing antibodies against either
KAT or MFP on the beads. The eluted antigens
were run on duplicate SDS-PAGE (4-12% Bis-
Tris, Invitrogen) and blotted onto nitrocellulose
membranes. Western blots were then developed
using polyclonal antibodies against either MFP or
KAT as primary antibodies. The MFP antibodies
were raised against recombinant AtMFP2 and the
KAT antibodies were raised against recombinant
Brassica napus KAT2 (48). The AtMFP2
antibodies have also been shown to cross-react
with recombinant AtAIM1. A control pull-down
was performed using immobilized antibodies
against His-tags with the peroxisomal lysate.
Structure analysis and modeling - Secondary
structure assignment and protein-protein
interactions were defined using PROMOTIF (49)
via the PDBSUM server
http://www.ebi.ac.uk/pdbsum/ (50). To construct
the Arabidopsis -oxidation complex model based
on the bacterial multi-enzyme -oxidation
complex, we simply superimposed the AtMFP2
structure (51) onto the PfMFP (chains A and B)
and the AtKAT2 dimer onto the PfKAT (chains C
and D; PDB code 1WDK) and then performed a
simulated annealing energy minimization in CNS
(52) in order to minimize clashes between side
chains of the two proteins, back-bone atoms
remained unchanged. To look at the amino acid
conservation of the prokaryotic PfKAT and
peroxisomal eukaryotic AtKAT2 their sequences
were subjected to a BLAST (53) search and then
unique sequences (56 for PfKAT (sequence
identity 57-99%) and 52 for AtKAT2 (sequence
identity 43-96%)) were compiled and aligned in
CLUSTALW (54). These multiple sequence
alignments were then used with their respective
structures in CONSURF (55), the output of which
allows coloring of the PDB files with respect to
conservation, from blue being conserved, through
the rainbow, to red being unconserved. The
PfKAT and AtKAT2 structures were not used
alone for CONSURF as many unrelated thiolases
were selected by the server. Instead, we opted to
select a range of prokaryotic KATs for the PfKAT
and peroxisomal eukaryotic KATs for the AtKAT2
in order to keep the comparison focused on the -
oxidation pathway. The PfMFP and AtMFP
structures were subjected to the basic CONSURF
50 sequences selection with PSI-BLAST E-value
cut-off of 0.001. Analysis of the protein-protein
interactions was aided by the use of PDBSUM and
PROTORP (56). Gap volume index values were
calculated with PROTORP and NOXclass (57),
where these values varied slightly, an average has
been reported. Figures were prepared using Pymol
(http://pymol.sourceforge.net/) and for clarity, for
the online supplemental movie, a missing loop,
containing W175, in the AtKAT2inactive structure
has been modeled in using Modeller 7v7 (58).
RESULTS
The crystal structures of AtKAT2 and HaKAT -
Crystal structures of Arabidopsis and Helianthus
3-ketoacyl-CoA thiolase have been solved in their
dimeric, open, reduced, active state to high
resolution (1.5 Å and 1.8 Å respectively). AtKAT2
consists of residues A: 46-448, B: 47-448 and
HaKAT consists of residues A: 47-438, B: 45-438.
Data collection, refinement and validation details
are given in Table 1. In order to facilitate an easy
reference system for the secondary structure
elements, the sequences of AtKAT2 and HaKAT
were aligned with sequences of other known
KAT2 structures, namely from Arabidopsis in the
inactive form (AtKAT2inactive, 2C7Y (21)), Homo
sapiens (HsKAT, 2IIK), Saccharomyces
cerevisiae (ScKAT, 1AFW, (19)) and
Pseudomonas fragi (PfKAT, 1WDK chains C and
D (16)). -strands and helices ( and 310) were
defined using PDBSUM (50), incorporated and
numbered according to domains (Supplemental
Fig. S1 online). The AtKAT2 and HaKAT
structures, reported here, are very similar and
exhibit the expected thiolase-like fold as described
for the ScKAT structure (19). Each subunit
consists of three domains (Fig. 1A); the N-domain
and C-domain are related by a pseudo two-fold
axis and have the same / tertiary fold consisting
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of a central five-stranded -sheet where the fifth
strand is anti-parallel with strands 1-4. Two -
helices cover the outer side of the -sheets and one
-helix from each domain is buried between the
sheets. The Loop domain (L-domain) is composed
of four -helices (H2, H3, H4 and H5), two 310-
helices (H1 and H6a) and a -hairpin (S1 and S2;
-hairpin 1) that extends away from the globular
protein by approx. 25 Å. The dimer (consisting of
subunits A and B) is formed by interactions
mainly involving the N-domains.
Subunit A superimposes onto subunit B
with an r.m.s.d. 0.3 Å over 403 C-atoms in the
AtKAT2 structure and with an r.m.s.d. of 0.4 Å
over 391 C-atoms in the HaKAT structure. As
expected there are deviations in the position and
quality of electron density at the very extremities
of the N- and C-termini. There is some deviation
in the orientation of -hairpin 1, by as much as 5
Å in AtKAT2, and small differences (~1 Å) in the
-helical region of the L-domain. Electron density
is of continuous high quality with only a few
exceptions: The loop from the N-domain to the L-
domain and the extremities of -hairpin 1 are of
poorer quality and here side chains are only
marginally defined. A mutation was also noticed
in the HaKAT electron density at position 223;
this was confirmed by sequencing to be a
threonine instead of an alanine. This residue is
situated on H4 in the L-domain and the mutation is
not thought to have affected the overall structure
since the AtKAT2 and HaKAT structures are
almost identical, as expected from their 85%
sequence identity, (dimer superimposes with an
r.m.s.d. 0.57Å over 780 C atoms). AtKAT2 and
HaKAT exhibit only small differences in two
loops; a movement of the -hairpin 1 in subunit B
that results in a C difference in position of ~7 Å
at the tip of the loop (Lys247AtKAT2 versus
Lys249HaKAT). This difference is only ~1 Å in A
subunits, suggesting a crystal packing influence.
To a much lesser extent there is also a variation of
the loop between the N-domain and the L-domain.
There is an additional -helix (H6a) in the C-
domain of AtKAT2 that is not observed in HaKAT
or in any of the other type I thiolase structures. In
addition to the side chains that are different
between these two proteins, a few changes in main
chain positions and side chain orientations are
observed, mainly in solvent exposed areas of the
structures. Fig 1B shows a superposition of all
type I thiolase structures available, these are
compared in more detail in the supplemental
material.
AtKAT2 redox potential and activity -
Sundaramoorthy and co-workers (2006) suggested
that the oxidized form found in their AtKAT2
crystals represented an inactive state of a
biologically relevant redox switch regulating
peroxisomal -oxidation in Arabidopsis. However
no data was presented bridging the form found in
their crystal structure to activity. Our activity data
(Fig. 2A) shows that AtKAT2 is inactivated, with
a half-life of 0.6 hours, in the presence of 10 mM
of the oxidant CSSC, while in the presence of 20
mM of the reductant CSH full activity is retained
for more than 12 hours (data not shown). In the
absence of both CSSC and CSH, AtKAT2 is
spontaneously inactivated with a half-life of 2.6
hours, indicating a very low reduction potential.
To investigate this further, tryptophan
fluorescence was recorded while titrating the
enzyme with CSH. Trp175 is the only tryptophan
residue in AtKAT2. Buried at the surface near the
active site and the subunit interface, it is optimally
positioned to report fluorescently on the redox-
coupled rearrangement of the molecule
(supplemental movie online). The tryptophan
fluorescence of AtKAT2 increases with increasing
[CSH]2/[CSSC] ratios (Fig. 2B). The increase is
accompanied by a small red-shift of the emission
(Fig. 2C) in accord with a slightly higher degree of
solvation of Trp175 in the open, active
conformation. Adding -mercaptoethanol to the
solution resulted in a similar increase, while H2O2
resulted in a decrease. Comparing the overall
secondary structure content of AtKAT2active and
the oxidized AtKAT2inactive there is a mere 2.4%
difference in the -helical content in the favor of
AtKAT2active, and 2.5% more -structure in the
AtKAT2inactive. CD-spectra of oxidized and reduced
AtKAT2 differ insignificantly, supporting the
interpretation that the fluorescence signals
represent the local rearrangements observed in the
AtKAT2inactive and AtKAT2active crystal structures
(please see supplemental movie online and
discussion). The transition midpoint between the
two states is 0.17+/-0.02 M, which by using the
Nernst equation, gives an Em at pH 8.5 of -372 mV
equal to E0 of -283 mV at pH 7.0 (assuming linear
proportionality with pH with slope -59 mV in this
pH range (40-41)), indicating that this redox
potential falls within a physiological range (see
discussion).
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Potential of KAT to form a multi-enzyme complex
with MFP - The existence of multi-functional
enzymes, which harbour the last three enzymatic
steps of -oxidation, in bacteria and mitochondria
led us to investigate whether a similar tri-
functional complex exists in plants. We employed
pull-down assays from enriched peroxisomal
lysate isolated from Brassica napus cv. seedlings
and focused on a possible interaction between
MFP2 and KAT2. Brassica and Arabidopsis both
belong to the mono-phyletic family of the
Brassicaceae and share a recent common ancestry
ending about 20 million years ago (59). The
BnKAT2 and AtKAT2 sequences are 96%
identical and the BnMFP2 and AtMFP2 sequences
are 90% identical. When BnKAT2 antibodies were
immobilized for co-immunoprecipitation, both
BnKAT2 and AtMFP2 antibodies identified bands
of an appropriate molecular weight on the
resulting western blot (see Fig. 3), suggesting that
these proteins interact. However, when AtMFP2
antibodies were immobilized, only the AtMFP2
antibodies identified a band on the western blot,
suggesting that KAT2 was not pulled down with
MFP2. The inability of the anti-AtMFP2 to co-
immunoprecipitate BnKAT2 does not discredit the
existence of the complex, as the antibodies may
disrupt the interaction. A negative control using an
unrelated antibody, anti-His, in place of the
antibodies against KAT2 and MFP2 for the pull-
down was performed and western blots were
developed using antibodies against KAT2 and
MFP2, no bands were identified (data not shown).
DISCUSSION
Redox regulation of AtKAT2 and comparison of
active and inactive structures - The redox
potential of the peroxisomes has not been
determined quantitatively, but the redox potential
in living plant cells has been determined to be
about -320 mV in the cytosol at pH 7.2 and -360
mV in the mitochondria at pH 8.0 (60). Assuming
that the peroxisomes can attain a similar redox
potential as the mitochondria, the ratio between
oxidized and reduced AtKAT2 at thermodynamic
equilibrium would be 0.24 at pH 8.5 while at -340
mV it would be 99% oxidized. Other dithiols that
have been suggested to be redox sensors, the
disulfides of Orp1 and the redox sensing domain
of the cytosolic Yap1 have potentials as low as -
315 and -330 mV at pH 7.0 (61), while the
potential of the active cysteines of wild-type plant
cytosolic thioredoxins varies from -280 to -304
mV at pH 7.0 (62). Yap1 is thought to be kept in
the reduced state by the non-equilibrium ratios of
102
– 104 reduced:oxidized thioredoxin existing in
the cells. Although a recent proteomics study
identified two putative thioredoxins and a
glutathione reductase in Arabidopsis peroxisomes
(63), there is little knowledge about disulfide
reducing agents in the peroxisomes. Taken
together, it is likely that AtKAT2 represents a very
sensitive redox switch capable of regulating -
oxidation even at minor perturbations of the
peroxisomal redox environment. Interestingly, it
has been observed that the activity of KAT was
compromised by 0.1 mM H2O2 in catalase
inhibited enzyme preparations from peroxisomes
of rat liver (64). It therefore seems likely that the
oxidized and reduced forms of AtKAT2,
crystallized under different conditions, represent
biologically relevant forms of AtKAT2.
When comparing the AtKAT2active (this
study) and AtKAT2inactive (pdb code: 2C7Y, (21))
structures, there are a number of differences. At
the monomer level Asp38-Asp51 in the
AtKAT2inactive structure form a -strand (S1a),
which affiliates with the C-terminal domain, and
pairs up with the equivalent strand in the dimer
formation (supplemental movie online) and results
in an extended central -sheet. -strand S1a is not
seen in the AtKAT2 structure described here,
despite the residues being present in the construct,
they are unseen in the electron density. One reason
for this difference could be the presence of the N-
terminal His6-tag in our construct, however we
think this is an unlikely explanation as the
equivalent residues are present in the HaKAT
construct and are also unseen in the structure
where there is no N-terminal His6-tag. The -
strand formation is therefore more likely the result
of the dimer reorientation in the oxidized form of
AtKAT2, caused by the disulphide bridge
formation. The inactive conformation provides a
different interface that allows the stabilization the
otherwise flexible chain, providing more
interaction. The active site loop in the inactive
form, between -strand S3 and -helix H5 in the
N-domain, is flipped out towards subunit B,
repositioning the Cys138 C by 5.4 Å
(superimposing A chains) or 5.8 Å (superimposing
B chains) away from the helix dipole that is
important for the activation of Cys138 (20). The
short 310-helix (N-domain H4) containing the
active site cysteine, present in all of the active
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structures, is not observed in the AtKAT2inactive
structure. The location of this loop-segment in
AtKAT2inactive would cause steric clashes with -
strand S3 in the N-domain and the preceding loop
on subunit B in the active dimer. As such, subunit
B is repositioned in the AtKAT2inactive dimer via a
rotation of ~30o and translation of ~10 Å
(supplemental movie online). The Met168-Pro195
region in AtKAT2active L-domain caps over the
S2a-H1 loop and -helix H3 in subunit B N-
domain and contains a short 310-helix (H1) and an
-helix (H2). In the AtKAT2inactive structure, this
loop is largely disordered, with Met173-Lys184
not seen in the structure, and residues Met168-
Pro172 extend out towards where -helix H1, in
the L-domain, is positioned in AtKAT2active. In
AtKAT2inactive, H2 and H3 in the L-domain form
one continuous -helix via a flip of Pro195 which
is then, remarkably, incorporated into the -helix
reorienting the polypeptide such that Cys192 is
repositioned by ~9.5 Å (supplemental movie
online). The large conformational change of this
region (C-terminus of N-domain S4 to N-terminus
of C-domain H4), upon the disulphide bridge
formation, results in the domain reorientation. -
hairpin 1 in the L-domain and S1-H1a in the C-
domain has a difference in position of ~6 Å and -
hairpin 2 has a difference of ~3 Å. When
superimposing the B subunits there is an
alternative movement in -hairpin 1; Lys249 C is
positioned 2.8 Å in the opposite direction to the 7
Å movement observed in this loop in the HaKAT
structure suggesting that this is highly mobile. The
310-helix, H6a (observed in AtKAT2active, HaKAT
and PfKAT), in the L-domain is not observed in
the inactive form; it is replaced with two -turns,
as in the HsKAT and ScKAT structures.
The AtKAT2inactive dimer is very different
from all other KAT structures and has roughly half
the amount of inter-subunit contacts as the active
dimers: 26 residues from each chain are involved,
an interacting surface area of 1225 Å2 and 11
hydrogen bonds and 168 non-bonded contacts
compared with 54 residues from each chain, an
interacting surface area of 2500 Å2, 41 hydrogen
bonds and 408 non-bonded contacts (values
calculated in PDBsum) for the AtKAT2active dimer.
In addition, the gap volume index (gives an
indication of how well the two protomers fit
together; a lower number indicates a better fit) is
3.9 Å for the inactive form and 2.3 Å for the active
form; and the inter-protomer salt bridges that are
present in all of the active KAT structures (Asp99-
Arg136 and Arg132-Asp147 (AtKAT2
numbering)) are not formed in the AtKAT2inactive
conformation. This strongly suggests that the
active, reduced conformation is the most stable
dimer form. If we take a look at the isoforms
present in Arabidopsis, both AtKAT1 and AtKAT5
possess the Cys192 and Pro195 and so are also
likely to be regulated via their redox environment.
In contrast, AtACAT1 and AtACAT2 (responsible
for the reverse reaction and share 34% and 36%
sequence identity with AtKAT2) do not have these
residues, suggesting that the reverse reaction
(Claisen condensation) is not regulated by redox.
Cys192 and Pro195 are conserved in all non-
mitochondrial eukaryotic KATs in which they
have been sought. However, these are absent in
the prokaryotic and mitochondrial KATs, thus
revealing a very distinct, perhaps
evolutionary/peroxisome versus mitochondria,
difference in the regulation of -oxidation.
Model of plant -oxidation multi-enzyme complex
- The crystal structures of the plant KATs
(presented here) and the crystal structure of
AtMFP2 (presented in the accompanying paper,
Arent et al. 2010) bear close resemblance in
structure to the components of the bacterial -
oxidation multi-enzyme complex from
Pseudomonas (PDB codes 1WDK, 1WDL,
1WDM and 2D3T (16)). The Pseudomonas
complex is composed of two MFP-subunits
(1WDK superimposes with AtMFP with r.m.s.d.
1.9 Å over 1406 residues, sequence identity 31%)
and two KAT-subunits (Fig. 4A) that constitute a
dimer that is indistinguishable to that of the
reduced plant AtKAT2active structure (1WDK
superimposes with r.m.s.d. 1.2 Å over 735 aa,
sequence identity 42%). The interactions between
the two MFPs and between the MFP and KAT
subunits in the Pseudomonas structure are quite
small in surface area, but collectively form a stable
complex that can withstand purification via gel
filtration (16,65). In addition, substrate dependent
conformational changes are seen within the
Pseudomonas tri-complex, mainly involving the
MFPs (compare PDB 1WDK with 2D3T for
extremities), but the protein:protein interactions
remain constant. Our co-immunoprecipitation
experiments suggest an interaction between MFP
and KAT in plants and thus, we were encouraged
to see if a complex, similar to the Pseudomonas
multi-enzyme complex, could be modeled from
our Arabidopsis KAT2 and MFP2 crystal
structures (see methods for details and Fig. 4). A
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complex between MFP2 and KAT2 would allow
metabolite channeling, facilitating the product of
one enzyme action to be transferred directly to the
next in the cycle, without either the equilibration
of the intermediate with the bulk solvent or the
pursual of an alternative pathway. Additionally,
the lack of substrate specificity at the final
thiolytic step of the pathway is consistent with the
idea of substrate channeling, while allowing the
redox control of the cycle as a whole in response
to H2O2 levels.
In the Pseudomonas structures, both MFP
subunits interact with both KAT subunits,
although the interacting surface areas are small
(500–850 Å2). There are two regions in the L-
domain of the KAT subunits that interact with
MFP subunits (Fig. 5): Region 1 is composed of
H1-H2 and has an interaction surface area of ~800
Å2 and a gap volume index of 2.5 Å indicating a
favorable complementary shape. Region 2
involves -hairpin 1 from the KAT subunit and
has an interaction surface area of ~550 Å2
and has
a gap volume index of 7 Å indicating a loose, less
complimentary shaped, interaction than that of
Region 1; there are few hydrogen bonds, some
favorable electrostatic interactions but no salt
bridges at this interface. The size and limited
specific interactions suggests that this complex
could be fairly transient in vivo. In region 1, H2
and the C-terminal of S1 are well conserved in the
prokaryotic KATs; however, H1 and the rest of the
-hairpin 1 seem to be fairly variable (Fig. 5).
Electrostatic interactions are evident between
Lys146 of H2 and the conserved negative surface
of the PfMFP (Asp162 and Glu166) (Fig. 5A, C).
Asp200, in the -hairpin, is in the vicinity of a
positively charged patch on the surface of the
MFP in the case of Pseudomonas. However, this
positive area is not at all conserved and in the
structures available there are no hydrogen bond
interactions observed. Asp135, in the loop
preceding H1, is actually situated near to a
negatively charged area of the MFP. Overall, the
interactions between region 1 in the KAT with the
MFP are more substantial, when compared with
region 2, with respect to surface area, conservation
and complimentary shape.
In comparison, the modeled complex for
AtKAT2 and AtMFP2 is fairly similar. The
calculated interaction surface area overall in the
Arabidopsis model is actually larger than that in
the Pseudomonas structure with region 1, the H1-
H2 region, involved in an interface of ~750 Å2 and
region 2, the -hairpin, involved in an interface of
~920 Å2. However, the shape fit of region 1 is not
as favorable as that in the Pseudomonas structure,
with a gap volume index of 4 Å. Region 2, which
is more extensive in the Arabidopsis model, has a
gap volume of 4.3 Å, which is more sustainable as
a contact than this region in the Pseudomonas
structure (7 Å). Residues are similarly conserved.
It should be noted that upon actual complex
formation small movements may occur providing
a different fit than our simple model. -hairpin 1 is
longer in AtKAT2 and thus has the potential to
make a larger interface with AtMFP2. Lys244 on
S1 is positioned nicely to form an electrostatic
interaction with the indented negative surface of
MFP2 (Asp185; Fig. 5D) and these residues are
conserved in plant KAT/MFPs. In the case of
Arabidopsis, there is even a potential for a salt
bridge formation between Glu253 on S2 and
His179 in AtMFP2. Although these residues are
not 100% conserved among plant MFP/KATs the
charges are more so with the glutamic acid well
conserved and histidine sometimes being replaced
by arginine.
There are a number of conserved proline
residues in the vicinity of the interface that could
be key to add structural rigidity to the loops
involved in the interaction with MFP2 (Fig. 5C,
D). The potential electrostatic interaction for
Lys146 seen in the Pseudomonas structure does
not exist in the case of the Arabidopsis model, as
there is no charged residue at that position and
equally the surface potential of AtMFP2 is not
negative in that area. Indeed, the AtMFP2 area
around H2 is largely neutral in surface potential
compared with the negative patch seen in
Pseudomonas. Overall, the Arabidopsis model
does exhibit more electrostatic potential (10-20%
more charged residues at interface) at each of the
interfaces than the Pseudomonas structure, which
could play a role in interaction specificity of the
isoforms in Arabidopsis and may make the
complex less tightly bound in conventional protein
extraction procedures. In addition, in plants there
is the potential to form cation-pi interactions
between a highly conserved Lys281 in AtKAT2
with Phe233 in AtMFP2 (conserved in plant
MFP2-like, but not in AIM1-like isoforms). Thus,
we propose that a similar interaction to that found
in Pseudomonas could exist between KAT2 and
MFP2 in the peroxisomal plant system, however
we would suggest that the -hairpin 1 (region 2)
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would play a more significant role in Arabidopsis
than it does in the Pseudomonas complex.
Interestingly, it is the H2 in region 1 of the
proposed Arabidopsis complex that hosts Cys192
that is involved in the disulphide bridge formation
with the active site cysteine in the redox regulation
(see previous section). The redox control in the
peroxisomal KATs could provide a reason why the
interaction with MFP2 in this H1-H2 region is not
as prominent as it is in the Pseudomonas structure,
as when the dimer undergoes conformational
change this area is affected the most and
consequently needs to be more malleable. When
the KAT dimer is inactivated, and is in the closed
conformation, this would either force a
rearrangement of the MFP:MFP or, most likely,
break the MFP:KAT complex altogether. In order
to make up for this region needing to be flexible,
the extended -hairpin in plant peroxisomal KATs
would allow for an additional, compensating
surface area for the interaction to achieve stability.
In conclusion, -oxidation of fatty acids is
a fundamental process that occurs in peroxisomes
and plays essential roles at all stages of a plants
life, from germination to mature plant. We have
demonstrated that the peroxisomal thiolytic
cleavage of the two carbon unit, the last step of a
-oxidation cycle, is controlled by a sensitive
redox switch that could be physiologically
relevant with regards to the environment of the
peroxisome. The direct communication between
the breakdown of fatty acids, with other cellular
activities such as the neutralization of toxic
substances, H2O2 signaling and stress responses is
of key importance for the viability of the cell, and
indeed the plant. The structures described here
allow for a comparison with 3-ketoacyl-CoA
thiolases from other species, a discussion on
substrate binding, and a complete description of
the large conformational change that occurs
between the active and inactive forms of the
thiolase. In addition, we have been able to model a
complex with the AtMFP2 structure that would
allow substrate channeling resulting in a highly
efficient metabolon in a reducing environment but
would be destroyed in an oxidative environment.
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FOOTNOTES
*The Danish National Science Research Council supported this project through DANSYNC with a post-
doctoral stipend to SA and supported synchrotron beam time. Beamline scientists at stations X12, DESY,
Hamburg and ID23, ESRF, France are thanked for technical assistance during data collection, as is
Annette Kure Andreassen for general laboratory assistance. We are also very grateful to Dr Anders
Brandt, Carlsberg Laboratory for the BnKAT antibody.
Atomic coordinates and structure factors for AtKAT2 and HaKAT have been deposited at the
protein databank, PDBe codes 2WU9 and 2WUA, respectively (http://www.ebi.ac.uk). 4Present address: School of Biochemistry and Immunology, Trinity College Dublin, Dublin 2, Ireland.
5Present address: Danisco Ltd., Edwin Rahrs Vej 38, 8220 Brabrand, Denmark.
FIGURE LEGENDS
Figure 1. The structure of KAT. A, Cartoon schematic of AtKAT2 monomer, N domain in dark blue, C
domain in green and L- domain in cyan, secondary structure elements are labeled. B, Superposition of
KATs in open conformation. R.m.s.d.’s and sequence identities given in Supplementary Table S1.
Detailed comparison of these structures is also included in the supplementary material.
Figure 2. Characterization of redox state dependent activity. A, AtKAT2 is inactive in the presence of
CSSC and active in the presence of CSH. The rates of acetoacetyl-CoA cleaving by AtKAT2 were
determined as a function of time and incubation with oxidant or reductant. 12.5 μM enzyme preparations
were incubated at 25°C in 35 mM Tris (pH 8.5), 0.1 M NaCl and 20 mM CSH (), 10 mM CSSC () or
buffer (). The effect of CSH () or CSSC () on the substrate is included for reference. The
incubations were sampled at the given time points by rapid mixing and 2500 times dilution with 100 μM
CoA, 50 μM acetoacetyl-CoA in a micro-cuvette and recording of absorbance at 233 nm. The final
concentrations of AtKAT2, CSH and CSSC in the reactions were 5, 8 and 4 nM respectively. B, AtKAT2
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changes tertiary structure upon incubation with CSSC. AtKAT2 was incubated with different ratios of
[CSH]2/[CSSC] at 25°C for 10 min and tryptophan emission spectra were recorded exciting the samples
at 285 nm and reading emission at 333 nm. The total concentration of CSH and CSSC was kept constant
at 125 mg/L. A two-state equation was fitted to the normalized (line) data and the midpoint of transition
determined. C, Fluorescence spectra of reduced (full line) and oxidized (dotted line) AtKAT2 at 1 and
0.03 M [CSH]2/[CSSC] respectively. Conditions as in B except emission recorded from 325-450 nm.
Figure 3. Western blot analysis from co-immunoprecipitation studies. Lanes contained the following:
(1) Elution 1 from MFP beads (2) Load onto MFP beads (3) Peroxisomal lysate (4) Load onto KAT beads
(5) Elution 3 from KAT beads (6) Elution 2 from KAT beads (7) Elution 1 from KAT beads (8)
Molecular weight marker. A, shows the western blot which was developed with primary antibody against
KAT. B, shows the western blot which was developed with primary antibody against MFE. A band is
seen for MFP from the KAT2 pull-down suggesting a complex (red circle). Elution steps were only
performed after extensive washing.
Figure 4. Construction of the Arabidopsis KAT2-MFP2 complex model, based on the prokaryotic
multi-enzyme complex (PDB code 1WDK). The Pseudomonas structure A, with MFP in medium gray
and KAT in light gray; Region 1 (-helical) and region 2 (-hairpin 1) are highlighted in black.
Arabidopsis model B, follows the same coloring scheme as A with MFP2 in medium gray and KAT2
dimer in light gray.
Figure 5. Comparison of prokaryotic multi-enzyme complex with model complex of AtKAT2 and
AtMFP2. A, and C, are of the Pseudomonas structure; B, and D, are of the Arabidopsis model. PfMFP
and AtMFP are shown as an electrostatic surface colored according to electrostatic potential (red, most
electronegative - white - blue, most electropositive) and PfKAT and AtKAT2 as a cartoon colored
(rainbow coloring) according to conservation among prokaryotic and plant KATs, respectively (blue,
most conserved - green - yellow - red, unconserved).
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TABLES
Table 1. X-ray data collection, processing and refinement statistics
Data Set HaKAT AtKAT2
Data collection
Collection site ID23, ESRF, France. X12, DESY, Germany.
Wavelength (Å) 0.9762 1.0
Space Group P212121 P21
Unit cell dimensions a=78.6Å b=100.7Å c=105.3Å a=60.8Å b= 86.7Å c= 72.7Å
=106.7o
Resolution (Å) 78.6 – 1.8 (1.9 – 1.8)a 34.6 – 1.5 (1.58 – 1.5)
a
Rmerge % 8.6 (51.9)a 7.8 (50.0)
a
bRpim % 3.8 (26.9)
a 3.1 (23.7)
a
Mean (I)/ sd mean (I) 18.0 (2.6)a 15.2 (2.6)
a
Completeness (%) 99.2 (94.8)a 97.2 (81.2)
a
No. of unique reflections 77995 112318
Multiplicity 8.2 (5.4)a 5.1 (4.3)
a
cWilson B factor (Å
2) 28.44 12.00
Refinement and quality
Number of non hydrogen protein
atoms
5875 6151
Number of water molecules 709 1134
Number of ethylene glycol
molecules
0 9
Rwork (%) 18.5 (24.4)f 14.9 (20.9)
f
Rfree (%) 21.6 (28.6)f 17.7 (21.9)
f
Std. dev.
Bond angles (o)
Bond lengths (Å)
1.020
0.006
1.088
0.006
Mean B (Å2)
Main chain (Å2)
Side chains (Å2)
Solvent (Å2)
33.4
29.8
34.3
44.6
14.3
9.9
14.0
27.5
Ramachandran plot
Most favored (%)
Additionally allowed (%)
Generously allowed (%)
Disallowed (%)
91.8
7.4
0.5g
0.3h
93.4
6.0
0.3g
0.3h
aNumbers in parentheses refer to outer resolution bin.
bMultiplicity weighted Rmerge (66).
cFrom Truncate
analysis (67). dPhenix.refine (47).
eProcheck
(68).
fHighest resolution bin.
gHaKAT Gln135 and AtKAT2
Gln137 adopt strained conformation in active site. hHaKAT Lys64 and AtKAT2 Lys66 adopt strained
conformation, in well-defined density, in tight turn.
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FIGURES
FIGURE 1. The structure of KAT
A
B
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FIGURE 2. Characterization of redox state dependent activity
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FIGURE 3. Western blot analysis from co-immunoprecipitation studies
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FIGURE 4. Construction of the Arabidopsis KAT2-MFP2 complex model, based on the
prokaryotic multi-enzyme complex (PDB code 1WDK)
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FIGURE 5. Comparison of prokaryotic multi-enzyme complex with model complex of AtKAT2 and
AtMFP2
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Valerie E. Pye, Caspar E. Christensen, James H. Dyer, Susan Arent and Anette Henriksensensitive redox switch
Peroxisomal plant 3-ketoacyl-coa thiolases structure and activity are regulated by a
published online May 12, 2010J. Biol. Chem.
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